Fluidic device for long-term explant culture and imaging

ABSTRACT

Zebrafish are a powerful model for investigating cardiac repair due to their unique regenerative abilities, scalability, and compatibility with many genetic tools. However, characterizing the regeneration process in live adult zebrafish hearts has proved challenging because adult fish are opaque and explanted hearts in conventional culture conditions experience rapid declines in morphology and physiology. To overcome these limitations, we fabricated a fluidic device for culturing explanted adult zebrafish hearts with constant media perfusion that is also compatible with live imaging. Unlike hearts cultured in dishes for one week, the morphology and calcium activity of hearts cultured in the device for one week were largely similar to freshly explanted hearts. We also cultured injured hearts in the device and used live imaging techniques to continuously record the revascularization process over several days, demonstrating how our device enables unprecedented visual access to the multi-day process of adult zebrafish heart regeneration.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application claims the benefit of U.S. provisional application Ser.No. 62/746,666 filed Oct. 17, 2018, the disclosure of which is herebyincorporated in its entirety by reference herein.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

This invention was made with government support under HL130172 awardedby National Institutes of Health/National Heart, Lung, and BloodInstitute. The government has certain rights in the invention.

TECHNICAL FIELD

In at least one aspect, the present invention is related to fluidicdevices for culturing and imaging organs.

BACKGROUND

Adult human myocardium has an extremely limited regenerative capacity(1). As a result, major cardiac injuries, such as myocardialinfarctions, are irreversible and often lead to severe long-termcomplications, including heart failure (2). Unlike humans, the adultmyocardium of several non-mammalian vertebrates can regenerate,including that of the zebrafish (3-7). The adult zebrafish heart canfully regenerate within two months after up to 20% of the ventricle isresected (8) largely due to the ability of pre-existing cardiac myocytesto undergo limited dedifferentiation followed by proliferation (9, 10).Many non-myocyte components of the myocardium, such as epicardial cells(11), the vasculature (12, 13), and the extracellular matrix (14), alsoplay important roles in regeneration. However, how the collectivecellular and acellular features of the heart interact and cooperate toaccomplish regeneration is incompletely understood. Establishing thesedetails is the first step towards potentially translating mechanisms ofzebrafish heart regeneration to human heart repair.

The existing gaps in knowledge related to adult zebrafish heartregeneration are in large part due to the limitations of existingexperimental tools to interrogate this process. Although epicardialcells (15) and cardiac myocytes (16, 17) from adult zebrafish heartshave successfully been isolated and cultured in vitro, these monolayercultures of a single cell type cannot be used to investigate cell-cellinteractions or organ-level mechanisms of regeneration. These types ofquestions are usually addressed by injuring hearts in vivo and employinghistological methods to evaluate features, such as wound size andcellular proliferation, at different time points after surgery assnapshots (11). However, this strategy provides limited dynamicinformation, such as pathways of cell migration or origins of differentcell types.

Because adult zebrafish are not transparent, imaging explanted hearts exvivo is the only option for imaging heart regeneration in real-time.Today, explanted hearts are usually cultured in Petri dishes ormulti-well plates. In these static conditions, hearts experience rapiddeclines in native-like morphology and function within three days (18),introducing artifacts that render them largely unusable for monitoringregeneration, a multi-day process. These morphological and functionaldeclines have been partially alleviated in a subset of hearts thatundergo gentle agitation in culture dishes (19). However, because thehearts are not confined to a specific location or orientation, agitationculture is not compatible with continuous live imaging, which isrequired to monitor processes that occur over several hours, such ascell migration.

In recent years, microfabricated fluidic devices have been implementedto culture arrays of zebrafish embryos and larvae for up to 72 hours,often with constant media perfusion to maintain viability (20-22).Within these devices, each organism is cultured within a compartmentslightly larger than the organism itself, which facilitates continuouslive imaging over several hours or days by minimizing arbitrary motionand preserving the orientation of the organism.

SUMMARY

In at least one aspect, the present invention provides an innovativefluidic chip system that holds juvenile or adult zebrafish hearts in asingle position, provides a continuous media flow, fits intoconventional optical and fluorescent imaging systems, and allows forpost-analysis specimen recovery. Unlike existing fluidic culturedevices, it focuses on supporting a specific organ for at least one weekwhile preserving organ structure and function in comparison to standardcell culture techniques. Produced using rapid prototyping and softlithography techniques, these devices are suitable for medium to highscale production and are easily customizable to other explanted organsor organoids that are 1-10 mm in diameter. The effectiveness of thisdevice for long-term explant culture is demonstrated by comparing thefunctional and histological characteristics of zebrafish hearts intraditional culture conditions and hearts in the chips with perfusion.The effectiveness of this device for live imaging is demonstrated byimaging the revascularization of an injured zebrafish heart maintainedin the device for 4.5 days.

BRIEF DESCRIPTION OF THE DRAWINGS

The patent or application file contains at least one drawing executed incolor. Copies of this patent or patent application publication withcolor drawing(s) will be provided by the Office upon request and paymentof the necessary fee.

For a further understanding of the nature, objects, and advantages ofthe present disclosure, reference should be had to the followingdetailed description, read in conjunction with the following drawings,wherein like reference numerals denote like elements and wherein:

FIGS. 1A, 1B, and 1C. Design of the zebrafish heart device. (A) Anexploded view of the entire device, including the center PDMS insertwith the triangular wells and channels, the PTFE membrane, thetransparent acrylic panels, the nuts and bolts, and the barbedconnectors. (B) A perspective view of the assembled device. (C) A topview of the perspective device.

FIGS. 2A, 2B, and 2C. (A) Perspective view of a triangular well. (B)Side view of a triangular well. (C) Top view of a triangular well.

FIG. 3 provides a perspective view showing the complete flow system ofthe microfluidic assembly.

FIGS. 4A and 4B provide a perspective view of molds used to form thePDMS inserts of the microfluidic assemblies of FIG. 1 .

FIGS. 5A, 5B, and 5C. (A) Photograph of the micro-milled polycarbonatetemplate used to mold the PDMS slab. (B) Photograph of a completeassembled device. (C) Simulated fluid flow streamlines (red) and surfaceshear stresses (color bar) through a complete device (top) and a sideview through a single well housing an average-sized heart (bottom).Scale bars are 10 mm for (A), and (B), and 1 mm for (C).

FIGS. 6A and 6B. Explanted zebrafish hearts in the fluidic device. (A)Photograph of PDMS slab immediately after loading with four explantedzebrafish hearts. Scale bar is 10 mm. (B) Fluorescent image ofcontracting heart explanted from fli1a:GFP transgenic zebrafish andimaged in the device. Image is a single frame from a movie of the hears.Scale bar is 200 μm.

FIGS. 7A, 7B, 7C, 7D, 7E, 7F, and 7G. Structural characterization ofFresh and one- and two-week Dish and Device hearts. (A-E) AFOG-stainedhistology samples of hearts from the indicated experimental groups, withhistology scores indicated. Scale bar is 200 μm. (F) Higher-resolutionimages of a sub-set of histology samples to illustrate increasing levelsof abnormalities and corresponding scores. Scale bars is 50 μm. (G)Histology scores (1-3) for all hearts. *p<0.05 and ** p<0.01 compared toFresh hearts, according to the Kruskal-Wallis test followed by Dunn'smultiple comparisons test to Fresh hearts. n=8, 8, 12, 8, and 9 forFresh, one-week Dish, one-week Device, two-week Dish, and two-weekDevice hearts, respectively.

FIGS. 8A, 8B, 8C, 8D, and 8E. Imaging and analysis of calcium activityin Fresh and one- and two-week Dish and Device hearts. (A) Image of aone-week Device heart incubated with Rhod-2 and paced at 1 Hz. Higherresolution images illustrate changes in Rhod-2 intensity on the edges ofthe heart at rest (left image) and at the peak of the transient (rightimage). Scale bar is 500 μm. (B) Representative calcium transients froma Fresh heart (black), one-week Dish heart (blue), and one-week Deviceheart (red), each paced at 1.0 Hz. The circled time points correspond tothe two smaller images in panel (A). (C) Capture rates for all hearts.*p<0.05 and **p<0.01 compared to Fresh hearts, according to theKruskal-Wallis test followed by Dunn's multiple comparisons test toFresh hearts. n=16, 10, 10, 11, and 12 for Fresh, one-week Dish,one-week Device, two-week Dish, and two-week Device hearts,respectively. (D) Rise and (E) decay times for all hearts. *p<0.05 and**p<0.01 compared to Fresh hearts, according to two-way ANOVA followedby Tukey's multiple comparisons test compared to Fresh hearts. n=10 forFresh hearts at all frequencies; n=6 for one-week Dish hearts at allfrequencies; n=9, 8, 7, and 8 for one-week Device hearts at 0.5, 1.0,1.5, and 2.0 Hz, respectively.

FIG. 9 . Zebrafish heart regeneration ex vivo. A zebrafish heartunderwent apex resection surgery followed by six days in vivo recovery.Live imaging of the apex wound site, demarcated with a dashed line (1),was then conducted on a six days post-amputation device-cultured adulttransgenic zebrafish heart expressing the endothelial marker fli1a:GFP(magenta) and flt1enh:tdTomato (cyan). This time-lapse imaging showsmigration of endothelial cells into the wound site (3) directly from theproximal vasculature. These cells form interconnections (6), followed bya visible restriction of the regenerated tissue around the wound site.Frames are selected from a movie of the hearts. Scale bar is 100 μm.

DETAILED DESCRIPTION

Reference will now be made in detail to presently preferred embodimentsand methods of the present invention, which constitute the best modes ofpracticing the invention presently known to the inventors. The Figuresare not necessarily to scale. However, it is to be understood that thedisclosed embodiments are merely exemplary of the invention that may beembodied in various and alternative forms. Therefore, specific detailsdisclosed herein are not to be interpreted as limiting, but merely as arepresentative basis for any aspect of the invention and/or as arepresentative basis for teaching one skilled in the art to variouslyemploy the present invention.

It is also to be understood that this invention is not limited to thespecific embodiments and methods described below, as specific componentsand/or conditions may, of course, vary. Furthermore, the terminologyused herein is used only for the purpose of describing particularembodiments of the present invention and is not intended to be limitingin any way.

It must also be noted that, as used in the specification and theappended claims, the singular form “a,” “an,” and “the” comprise pluralreferents unless the context clearly indicates otherwise. For example,reference to a component in the singular is intended to comprise aplurality of components.

The term “comprising” is synonymous with “including,” “having.”“containing,” or “characterized by.” These terms are inclusive andopen-ended and do not exclude additional, unrecited elements or methodsteps.

The phrase “consisting of” excludes any element, step, or ingredient notspecified in the claim. When this phrase appears in a clause of the bodyof a claim, rather than immediately following the preamble, it limitsonly the element set forth in that clause; other elements are notexcluded from the claim as a whole.

The phrase “consisting essentially of” limits the scope of a claim tothe specified materials or steps, plus those that do not materiallyaffect the basic and novel characteristic(s) of the claimed subjectmatter.

With respect to the terms “comprising,” “consisting of,” and “consistingessentially of,” where one of these three terms is used herein, thepresently disclosed and claimed subject matter can include the use ofeither of the other two terms.

It should also be appreciated that integer ranges explicitly include allintervening integers. For example, the integer range 1-10 explicitlyincludes 1, 2, 3, 4, 5, 6, 7, 8, 9, and 10. Similarly, the range 1 to100 includes 1, 2, 3, 4 . . . 97, 98, 99, 100. Similarly, when any rangeis called for, intervening numbers that are increments of the differencebetween the upper limit and the lower limit divided by 10 can be takenas alternative upper or lower limits. For example, if the range is 1.1to 2.1 the following numbers 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, and2.0 can be selected as lower or upper limits.

In the examples set forth herein, concentrations, temperature, andreaction conditions (e.g., pressure, pH, flow rates, etc.) can bepracticed with plus or minus 50 percent of the values indicated roundedto or truncated to two significant figures of the value provided in theexamples. In a refinement, concentrations, temperature, and reactionconditions (e.g., pressure, pH, flow rates, etc.) can be practiced withplus or minus 30 percent of the values indicated rounded to or truncatedto two significant figures of the value provided in the examples. Inanother refinement, concentrations, temperature, and reaction conditions(e.g., pressure, pH, flow rates, etc.) can be practiced with plus orminus 10 percent of the values indicated rounded to or truncated to twosignificant figures of the value provided in the examples.

For any device described herein, linear dimensions and angles can beconstructed with plus or minus 50 percent of the values indicatedrounded to or truncated to two significant figures of the value providedin the examples. In a refinement, linear dimensions and angles can beconstructed with plus or minus 30 percent of the values indicatedrounded to or truncated to two significant figures of the value providedin the examples. In another refinement, linear dimensions and angles canbe constructed with plus or minus 10 percent of the values indicatedrounded to or truncated to two significant figures of the value providedin the examples.

Throughout this application, where publications are referenced, thedisclosures of these publications in their entireties are herebyincorporated by reference into this application to more fully describethe state of the art to which this invention pertains.

Abbreviations

“PDMS” means polydimethylsiloxane.

“PTFE” means polytetrafluoroethylene.

With reference to FIGS. 1A, 1B, 1C, 2A, 2B, 2C, and 3 , schematicillustrations of a microfluidic device for imaging whole organs isprovided. The microfluidic device is typically transparent at positionsthat allow imaging of organs, organoids, or organisms placed therein.Moreover, microfluidic assembly allows cultured medium to nourishexplanted organs placed therein. Microfluidic assembly 10 includespolymeric block 12 having a top surface 14 and a bottom surface 16.Typically, polymeric block 12 is made from polymeric organosilicons suchas PDMS. Polymeric block 12 defines an input reservoir 18 and an outputreservoir 20. Typically, input reservoir 18 and output reservoir 20extend from top surface 14 to approximately 0.5 mm from the bottomsurface 16. Polymer block 12 also defines at least one triangular well24 which extends from top surface 14 to bottom surface 16. Typically,polymeric bock 12 has a thickness from 1 mm to 10 mm. Triangular well 24traps and maintains the orientation of explanted organs (e.g., zebrafishhearts), organoids, or organisms. The well is triangular in the sense ofincluding sidewalls 26 and 27 which are angled with respect to eachother such that these sidewalls approach each other along direction d₁.Direction d₁, is generally in the average direction of the liquid flow.The convergence of sidewalls 26 and 27 allow a specimen to be wedged inplace thereby fixed its position and orientation. Advantageously, themicrofluidic assembly can be used to culture mouse embryos/organs andhuman fetal organs. In a refinement, polymer block 12 defines aplurality of triangular wells 24 (e.g., 2 to 10) which allow multipleorganoids, or organisms, to be studied in parallel.

Still referring to FIGS. 1A, 1B, 1C, 2A, 2B, 2C, and 3 , polymer block12 also defines first flow channel system 28 and second flow channelsystem 30. First flow channel system 28 is in fluid communication withthe input reservoir 18 and the at least one triangular well. Similarly,second flow channel system 30 is in fluid communication with the outputreservoir 20 and the at least one triangular well. Each of first flowchannel system 28 and second flow channel system 30 include tracks thathave a closed bottom and open top. Typically, each track in first flowchannel system 28 and second flow channel system 30 have a depth fromabout 300 to 800 μm and a width from about 300 to 800 μm. The tracks offirst flow channel system 28 and second flow channel system 30 enter andexit respectively triangular wells 24 at top surface 14. Moreover, theflow of media is uniformly distributed across first flow channel system28 and second flow channel system 30, which are designed to be equal inhydraulic resistance as the media is perfused microfluidic assembly 10.Finally, polymer block 12 defines inlet conduit 34 in fluidcommunication with the input reservoir 18 and outlet conduit 36 in fluidcommunication with the output reservoir 20. Flow adapters/connectors 38,39 can be placed in inlet conduit 34 and outlet conduit 36 to allow flowof a liquid media into and out of microfluidic assembly 10.

First transparent plate 40 is disposed over the top surface 14 of thepolymeric block 12 and second transparent plate 42 disposed over thebottom surface 16 of the polymeric block. In a refinement, gas permeablemembrane 48 is interposed between the first transparent plate 40 and thepolymeric block 42. Gas permeable membrane 48 has several openings 49that allow imaging of each of the triangular wells 24. In a refinement,gas permeable membrane 48 is a PTFE membrane. In a refinement, firsttransparent plate 40 defines a first vent slot 50 and a second vent slot52 allowing gases to vent from the gas permeable membrane 48. First ventslot 50 overlays the input reservoir 18 and a portion of first flowchannel system 28, and the second vent slot 52 overlays the outputreservoir 20 and a portion of second flow channel system 30. Finally,polymeric block 12, first transparent plate 40, second transparent plate42, and gas permeable membrane 48 are bolted together with bolts 56 andnuts 58 via suitable access holes in each of these components.

With reference to FIGS. 2A, 2B, and 2C, triangular well 24 is defined byconverging sidewalls 26 and 27, upstream sidewall 60, and downstreamsidewall 62, top wall 64, and bottom wall 66. Top wall 64 is defined byfirst transparent plate 40 while bottom wall 66 is defined by secondtransparent plate 42. Sidewalls 26 and 27 are generally planar and areangled with respect to each other such that these sidewalls approacheach other along direction d₁. The imaginary extension of sidewalls 26and 27 intersect at imaginary line l₁. The angle a₁ defined in a planeperpendicular to sidewalls 26 and 27 is an angle typically between 10and 120 degrees. In a refinement, angle a₁ is from about between 30 and90 degrees. Liquid flows into triangular well 24 via flow channel 70which is part of first flow channel system 28. In a refinement, flowchannel 70 enters triangular well 24 through an opening in upstreamsidewall 60. Similarly, liquid flow out of triangular well 24 via flowchannel 72 which is part of second flow channel system 30. In arefinement, flow channel 72 exists triangular well 24 through an openingin downstream sidewall 62. In a further refinement, each of flowchannels 70 and 72 are positioned at the top of triangular well 24.

Referring to FIG. 3 , a schematic of the flow channeling system definedby microfluidic assembly 10 is provided. Fluid is introduced into inletconduit 34 and through each of triangular wells 24. An explanted organ80 (e.g., zebrafish heart) is placed at the bottom of each of triangularwells 24. The triangular cross-section of the well maintains theorientation of the explanted organ during imaging.

In another embodiment, a method for imaging an explanted organ applyingthe microfluidic assembly set forth above is provided. The methodincludes a step of placing an explanted organ into the at least onetriangular well and then continuingly flowing a culture medium into themicrofluidic assembly. The explanted organ is imaged according totechniques known to those skilled in the art of organ culturing andimaging. As set forth above, microfluidic assembly 10 can include aplurality of triangular wells with an explanted organ (e.g., a zebrafishheart) being placed into each triangular well. Typically, the culturemedium includes a fluorescent dye or indicator which enhances imaging ofthe organ.

In another embodiment, a method for making the microfluidic assembly ofFIGS. 1-3 are provided. FIG. 4A provides a perspective view of a mold 84that can be used to form a device with a single triangular well whileFIG. 4B provides a perspective view of a mold 86 that can be used toform device with a plurality (e.g., 4) triangular wells. The methodincludes a step of providing a mold (e.g., mold 84 or 86) havingsidewalls that define a mold cavity 88. Characteristically, the moldcavity includes protrusion 90 for forming an input reservoir, protrusion92 for forming at least one triangular well, protrusion 94 for formingan output reservoir, protrusion 96 for forming a first flow channel influid communication with the input reservoir and the at least onetriangular well, and protrusion 98 for forming a second flow channelsystem in fluid communication with the output reservoir and the at leastone triangular well. A curable resin is introduced into the mold cavityand then cured or allowed to cure. A polymeric block having the featureset forth above is removed from the mold cavity. The polymeric block ispositioned between a first transparent plate and a second transparentplate. In a refinement, a gas-permeable membrane as set forth above isplaced between the first transparent plate and a second transparentplate.

The following examples illustrate the various embodiments of the presentinvention. Those skilled in the art will recognize many variations thatare within the spirit of the present invention and scope of the claims.

A variety of rapid prototyping and soft lithography techniques were usedto fabricate a fluidic device that can house up to four adult zebrafishhearts in parallel with continuous media perfusion. To ensurecompatibility with live imaging, the device confines each heart to atriangular chamber slightly larger than an average heart and the bottomof the device is an optically-clear acrylic panel. The device is alsoreversibly assembled so that hearts can be inserted and later removedfor additional characterization, such as histology or gene or proteinexpression analysis. When cultured in the device, explanted adultzebrafish hearts experienced significantly fewer structural andfunctional declines compared to hearts cultured in Petri dishes over aone-week period. We also implemented our device to culture an injuredadult zebrafish heart and successfully monitored regeneration usingcontinuous live imaging over 4.5 days. Thus, our fluidic device enablesunprecedented access to adult zebrafish heart regeneration, which canadvance our fundamental knowledge of this process and potentially leadto the identification of molecules or processes that can translate tohuman heart repair.

Design and Fabrication of the Fluidic Device

We designed a fluidic device (FIG. 1A) for culturing explanted adultzebrafish hearts that has the following components: (1) a PDMS slabembedded with a fluid path comprising inlet and outlet media reservoirs,a network of channels, and four triangular compartments that areslightly larger than an average adult zebrafish heart; (2) adaptors forconnecting inlet and outlet reservoirs to tubing and a syringe pump formedia perfusion; (3) a PTFE membrane for capturing and releasing airbubbles; (4) two transparent acrylic panels that compress the PDMS slaband PTFE membrane to enclose the fluid path; and (5) nuts and bolts forreversible device assembly. To fabricate the device, we milled theinverse of the fluid path into a piece of polycarbonate (FIG. 4B, FIG.5A), which served as a master template for molding slabs of PDMSembedded with the fluid path. For each PDMS slab, we accessed the fluidpath by punching inlet and output ports and inserting and sealing barbednylon connectors that can interface with tubing. Next, we laser-cut thetop and bottom panels from transparent 1.5 mm-thick acrylic sheets,which are thin enough to fit within the working distance of mostlow-numerical aperture or long working distance microscope objectives.We also laser-cut PTFE membranes and inserted one between each PDMS slaband top acrylic panel. Because PTFE is hydrophobic and gas-permeable,gas bubbles accumulate on PTFE and escape through the membrane underpressure (23). To vent the air bubbles collected by the PTFE membrane,we laser-cut rectangular voids into the top acrylic panel above theinlet and outlet channels. Because PTFE is opaque, we also laser-cuttriangular holes in the PTFE membrane aligned above the triangular wellsto avoid obstructing the imaging of the hearts.

At the corners of the PDMS, PTFE membrane, and acrylic panels, wepunched or laser-cut holes to accommodate bolts. When bolted between thetwo panels of acrylic and PTFE membrane, the elastomeric PDMS acts as agasket and forms a liquid-tight seal around the wells and channels.However, because the components are not permanently bonded to eachother, the device can be easily assembled and disassembled so thatintact hearts can be placed in the device and then later retrieved fordownstream analyses, such as histology or calcium imaging. The completeassembled device (FIG. 1B, FIG. 1C, FIG. 5B) has a footprint of 20 mm×20mm to be compatible with most standard microscope stages.

Simulation of Flow and Shear Stresses in the Device

To perfuse media through our device, we chose a flow rate of 250 μL/hrbecause that is near the lower limit of most standard 60 mL syringepumps and therefore was a practical choice for minimizing shear stress.To calculate shear stresses experienced by average zebrafish heartscultured in the device at this flow rate, we used multi-physics modelingsoftware. As shown by the fluid streamlines in FIG. 5C, our simulationspredict that media is perfused throughout each compartment and thatshear stresses reach a maximum of approximately 250 μPa on the topsurface of the heart. This is well below the reported 1-3 Pa maximumshear stress applied to zebrafish heart walls during normal beating (24,25), ensuring that hearts in the device do not experiencesupraphysiological shear stresses that may induce injury.

Because the total volume of the complete fluid path is approximately 78μL, the media is completely refreshed approximately three times perhour. At this flow rate, the syringe pump can operate continuously forten days without user intervention when using a full 60 mL syringe. Thisflow rate is also adequate for nutrient exchange because the standardpractice for culturing explanted hearts is to exchange 3-5 mL of mediaevery 1-2 days, which equates to approximately 60-200 μL/hr. Thus, amedia perfusion rate of 250 μL/hr ensures that hearts have adequatemedia exchange without experiencing any injury from shear stress.

Structural Characterization of Cultured Hearts

To evaluate the live imaging capabilities of our device, we nextexplanted hearts from adult transgenic zebrafish expressing thepan-endothelial marker fli1a:GFP (26) and placed them in a device toimage the ventral aspect of the ventricle such that their outflow tractswere in-line with outlet channels (FIG. 6A). Because hearts retain somespontaneous beating, this orientation ensures that any heart-generatedflow through the lumen would be in alignment with pump-generated flow.We then connected the assembled device to a syringe pump, mounted thedevice on a fluorescent microscope, and initiated media perfusion. Witha low-power objective, we could image through the bottom acrylic paneland record the GFP signal as the heart contracted (FIG. 6B). Because theheart was confined to a triangular compartment, its position stayedconstant throughout the acquisition. This feature enables morphologicalfeatures, such as changes in the geometry of the vascular network, to beeasily tracked over multiple cardiac cycles.

To determine if culturing hearts in the device extended their lifetimein culture, we next explanted hearts from adult wildtype zebrafish,loaded four hearts per device, connected each assembled device to asyringe pump, placed them in an incubator, and initiated mediaperfusion. At one- and two-week timepoints, we removed hearts from thedevice for histological staining. We also performed similar staining onfreshly explanted hearts and hearts cultured in conventional Petridishes for one and two week(s). In Fresh hearts, the myocardium wascompact throughout the cortical and trabecular layers in all hearts(FIG. 7A), as expected. One-week Dish hearts showed varying amounts ofstructural degradation, including graining and discoloration of themyocardium (FIG. 7B). In most of these hearts, morphological changeswere primarily localized to the cortical layer (FIG. 7Ba), but didextend into the trabecular layer in two of the eight hearts (FIG. 7Bb).In contrast, only two of the twelve one-week Device hearts showedabnormalities (FIG. 7C). Unlike the Dish hearts, these abnormalitiesappeared as a separation of the cortical and trabecular layers ratherthan a decrease in the integrity of the tissue itself, which stillappeared relatively healthy (FIG. 7Cb). The remainder of the one-weekDevice hearts showed almost no signs of degradation and appeared similarto Fresh hearts (FIG. 7Ca). None of the one-week Device hearts showedany abnormalities in the inner trabecular layer of the myocardium. Aftertwo weeks, half of the Device hearts (FIG. 7D) and all Dish hearts (FIG.7E) showed abnormal trabecular myocardium. Additionally, half of theDish hearts presented with ectopic collagen staining within themyocardium.

To quantify these observations, we scored the hearts from 1-3 based onthe severity and positioning of any abnormal morphologies. The score wasbased on two descriptive categories: cortical abnormality or degradation(any abnormal morphology restricted to the outer layer of cardiactissue), and extensive abnormality (any abnormal morphology ordegradation that extends into the inner trabecular layer of cardiactissue). Examples of abnormal morphologies and corresponding scores areshown in FIG. 7F. Combined histology scores (FIG. 7G) indicate thatone-week Dish hearts had a significantly worse score than Fresh hearts(p=0.0115) whereas one-week Device hearts were not statisticallydifferent than Fresh hearts (p>0.9999). After two weeks, both Dish andDevice hearts were significantly different than Fresh hearts (p<0.0001and p=0.0007, respectively). These results suggest that culturing heartsin the fluidic device delayed degenerative processes observed inconventional culture conditions.

Functional Characterization of Cultured Hearts

To characterize the functional properties of cultured hearts, wemeasured calcium activity in Fresh and one- and two-week Dish and Devicehearts while pacing hearts at 0.5, 1.0, 1.5, and 2.0 Hz (FIG. 8A-B).First, we quantified the capture rate at each frequency (FIG. 8C),defined as the percentage of detectable calcium transients perstimulation pulse. As expected, Fresh hearts had a capture rate of100±0% (n=9) at all four frequencies. One-week Dish hearts had capturerates of 80±42%, 41±38%, 25±33%, and 19±32% (n=10 for each) at 0.5, 1.0,1.5, and 2.0 Hz, respectively. These values were significantly lowerthan Fresh hearts at 1.0, 1.5, and 2.0 Hz (p=0.0047, p=0.0007, andp=0.0005, respectively), indicative of pronounced physiological declinein Dish hearts. In contrast, one-week Device hearts had capture rates of100±0%, 90±21%, 76±32%, and 63±34% (n=10 for each) at 0.5, 1.0, 1.5, and2.0 Hz, respectively. These values were significantly similar to Freshhearts at all frequences (p>0.9999 at 0.5, 1.0, and 1.5 Hz and p=0.3205at 2.0 Hz). These data suggest that Device hearts experienced fewerfunctional declines compared to Dish hearts after a one-week cultureperiod.

In addition to differences in capture rates, calcium transients fromone-week Dish hearts appeared wider than Fresh hearts and one-weekDevice hearts (FIG. 8B). To quantify this, we calculated calciumtransient rise (FIG. 8D) and decay (FIG. 8E) times. Because we can onlyanalyze transients from hearts that responded to pacing, thesecomparisons are a subset of the data shown in FIG. 8C. At allstimulation frequencies, rise and decay times were significantly longerin Dish hearts compared to Fresh hearts (p=0.0083, p=0.0003, p<0.0001,and p<0.0001 for rise time at 0.5, 1.0, 1.5, and 2.0 Hz; p<0.0001 fordecay times at 0.5, 1.0, 1.5, and 2.0 Hz). In contrast, the rise timefor Device hearts was significantly longer than Fresh hearts only at 1.5and 2.0 Hz (p=0.1711, p=0.0858, p=0.0241, and p=0.0164 at 0.5, 1.0, 1.5,and 2.0 Hz). Furthermore, the decay time for Device hearts wassignificantly longer than Fresh hearts only at 0.5 Hz (p=0.0052,p=0.1866, p=0.2108, and p=0.1220 at 0.5, 1.0, 1.5, and 2.0 Hz). Thus, ingeneral, calcium transient dynamics in one-week Device hearts were moresimilar to Fresh hearts compared to one-week Dish hearts.

After two weeks of culture, most of the two-week Dish hearts wereunresponsive to electrical stimulation, with average capture rates of7±13%, 2±4%, 2±4%, and 1±2% at 0.5, 1.0, 1.5, 2.0 Hz, respectively (n=12for each). In contrast, two-week Device hearts had capture rates of45±50%, 39±47%, 38±48%, and 34±49% at these same frequencies (n=12 foreach). Although two-week Device hearts were generally more responsivethan two-week Dish hearts, the capture rates for both groups weresignificantly lower than Fresh hearts (p<0.01 for all comparisons),likely due in part to the wide variability in responses.

Live Imaging of Heart Regeneration Ex Vivo

We next asked if adult zebrafish hearts maintain their regenerativecapacity when cultured in the device and, if so, if we could visualizethis process. To test this, we amputated zebrafish hearts expressing thepan-endothelial marker fli1a:GFP (26) and arterial specific markerflt1^(enh):tdTomato (27) and allowed initial responses to the injury toproceed in vive for six days. After the initial immune and wound healingresponses subsided, we explanted the hearts, inserted them in the devicesuch that the apex was at the base of the wells and bulbus arteriosus atthe top so that we could observe regeneration of the vasculature, andcollected an image every hour over the course of 4.5 days (FIG. 9 ).During this time, we observed the active migration of endothelial cellsinto the wound site from both coronary and non-arterial vessels proximalto the wound site between six and ten days post-amputation. It appearsthat most, if not all, vasculature was derived from this angiogenicmigration from pre-existing proximal vasculature. We also observed thatendothelial cells preceded, or migrated with, a progressive constrictionaround the wound site (FIG. 9-3 ). This may indicate that cellsbranching off the coronary vessels constrict the regenerating tissuearound the wound site, which could be coupled to other regenerativeprocesses. After projecting sprouts extended into the wound site, theendothelial cells formed interconnections with each other to generatethe dense plexus (FIG. 9-6 ). Collectively, these data demonstrate thatour device enables the visualization of regeneration in intact adultzebrafish hearts ex vivo, a process that has previously beeninaccessible to live imaging.

Discussion

In distinct contrast to adult human hearts, adult zebrafish hearts arehighly regenerative and can fully repair after severe injuries due tothe orchestration of multiple cellular and extracellular components(5-14). Mechanisms driving zebrafish heart regeneration couldpotentially translate to therapeutics that restore the function ofinjured human hearts, but only after the details of this process areestablished. However, the in vitro (15-17) and ex vivo (18, 19)approaches that exist today provide limited insight into mechanisms ofheart regeneration, especially the role of distinct cell types and theirinteractions after injury. To overcome these limitations, we fabricateda fluidic device for culturing explanted adult zebrafish hearts thatcontinuously perfuses the hearts with media and immobilizes them tofacilitate live imaging over several days. As shown by our histology andcalcium transient data, our fluidic device extended the culture lifetimeof explanted adult zebrafish hearts to one week in most cases and twoweeks in some cases. Furthermore, we cultured an injured heart withinour device and, over the course of 4.5 days, captured endothelial cellssprouting from existing vessels, migrating into a myocardial wound site,forming interconnections, and assembling a new vascular network thatlikely influences the migration and regeneration of other cell types.Previous lineage tracing data suggested that the likely source of thisvasculature was existing endothelial cells, but could not distinguishbetween the existing coronary vasculature or the underlying endocardium(28). Confocal imaging of fixed cryoinjured hearts appeared to showactively migrating vasculature on the edge of the woundsite andsuggested that revascularization is required for regeneration (12). Ourobservations confirm that this is the case, and further indicate thatboth arterial and non-arterial endothelial cells contribute to thisregenerating population and appear to migrate in a coordinated fashion.Thus, our device enables unprecedented access to dynamic mechanisms ofregeneration in intact adult zebrafish hearts, including cell migration,the origins of specialized tissues, and interactions between cardiaccell types. This approach can be widely adapted as a new experimentaltest-bed for establishing mechanisms of heart regeneration acrossmolecular, cellular, and organ-level scales, especially becausezebrafish are highly compatible with genomic modifications (29).

To design and fabricate our device, we used standard CAD software,microfabrication equipment, and low-cost materials. Thus, it isrelatively straightforward to re-configure our device to accommodateother explanted organs or tissues of similar scale, including neonatalmouse hearts, which are also regenerative (30), or organoids, which canmodel a variety of developmental, physiological, and pathologicalprocesses (31, 32). These types of mm-scale, 3-dimensional tissueconstructs could similarly benefit from the major features of ourfluidic device: (1) intact tissues can be inserted and removedon-demand, (2) media is continuously perfused, and (3) live-imaging overlong-term culture is feasible. Thus, our basic device framework has manyapplications for model systems beyond adult zebrafish hearts. However,one limitation of our device is that it is not compatible with highresolution imaging because the bottom panel is not glass, which is toobrittle for our current design. We hope to address this problem in afuture iteration of the device to enable dynamic imaging of processes oncellular and sub-cellular scales.

Intact zebrafish are currently being employed to screen the phenotypiceffects of drugs because they have a rapid growth rate and arerelatively easy and inexpensive to maintain, especially compared torodents (33). For these same reasons, culturing explanted zebrafishhearts in our fluidic device could be implemented as a new tool forefficiently screening the safety and/or efficacy of drugs on the heartex vivo. Although there are anatomical differences between zebrafishhearts and human hearts, they have many similarities in terms of theirelectrophysiology. For example, action potentials recorded fromzebrafish cardiac myocytes have a relatively long duration and plateauphase (34, 35), which matches human cardiac myocytes more closely thanmouse cardiac myocytes. This is especially advantageous for screeningthe arrhythmogenic effects of drugs (36). Additionally, due to theirease of genetic manipulation, mutations associated with arrhythmogenicdiseases, such as Long QT Syndrome (37), can be easily introduced intozebrafish (38). Thus, hearts from transgenic zebrafish with humandisease-relevant mutations could be explanted, cultured in our device,and used for live imaging and/or medium-throughput screening toefficiently identify potential therapeutic targets and molecules.Importantly, calcium transients recorded from hearts cultured in ourfluidic device for one week were more similar to freshly isolated heartscompared to hearts cultured in conventional dishes. Thus, our devicehelped preserve some of the electrophysiological features of hearts inculture, improving their relevance for screening the arrhythmic oranti-arrhythmic effects of drugs. However, a limitation of our device isthat it currently can only accommodate four hearts. Thus, our devicewould need to be modified to accommodate many more hearts to be usefulfor medium-throughput drug screening, which should be feasible withsimple design modifications.

Materials and Methods

Experimental Design

The objective of this study was to fabricate a fluidic device forculturing explanted zebrafish hearts that extends their lifetime inculture and accommodates live imaging over several days. To evaluate theperformance of our device, we implemented histology and calcium imagingtechniques to quantify the structure and function, respectively, ofhearts from the following experimental groups and endpoints: freshlyexplanted hearts (“Fresh hearts”), hearts cultured in the fluidic devicefor one or two weeks (“one-week Device hearts”, “two-week Devicehearts”), and hearts cultured in Petri dishes for one or two weeks(“one-week Dish hearts”, “two-week Dish hearts”). Each experimentalgroup consisted of 10-15 hearts, a sample size that is consistent withsimilar studies in this field. The study was not blinded and all datawas included (i.e., outliers were not defined or excluded). Forregeneration studies, hearts from previously characterized transgenicfish were used to visualize the revascularization of the wound area. Weoptimized the imaging parameters based on cellular toxicity and thespeed of endothelial cell migration. This allowed us to live image theprocess of regeneration and observe the contribution of proximalvascular endothelial cells in 3 out of 4 replicates.

Fluidic Device Design and Fabrication

The fluid path (FIGS. 1-3 ) was designed using SolidWorks 2018 CADPackage (Dassault Systèmes SolidWorks Corporation, Waltham, Mass., USA)and consists of four parallel channels (0.5 mm×0.5 mm) that lead to arecessed triangular compartment (depth: 3.5 mm, length: 3.5 mm, basewidth: 1.75 mm). The four channels branch from a single inlet reservoirand merge into a single outlet reservoir with the same dimensions(radius: 2 mm, depth: 3 mm). For all channels and wells, sharp anglesand intersections were rounded with fillets to minimize turbulence andair bubble accumulation. The fluid path was designed to fit within a 20mm×20 mm slab.

The inverse of the fluid path and slab was milled from polycarbonatestock material using an Othermill V2 micro-milling machine (OtherMachine Co, Berkeley, Calif., USA). All toolpaths were converted intog-code in Fusion360 (Autodesk, San Rafael, Calif., USA) and run usingOtherplan (Other Machine Co, Berkeley, Calif., USA). The clearingstrategy was based on a combination of adaptive roughing and horizontaland contour finishing using ⅛″, 1/16″, and 1/32″ flat end mills. Therecommended tool settings based on polycarbonate clearing for spindlespeed, cutting feed rate, plunge feed rate, and maximum stepdown sizewere taken from the manufacturer's website based on tool bit size. Thispolycarbonate template was vapor polished using previously describedmethods (39).

The polycarbonate template was used as a template for molding slabs ofpolydimethylsiloxane (PDMS, Sylgard 184; Dow Corning Corporation,Midland, Mich., USA). PDMS was combined at a 10:1 (w/w) ratio ofelastomer base to curing agent, mixed and degassed using a planetarycentrifugal Thinky Mixer AR-100 (Thinky Corporation, Tokyo, Japan),poured into the polycarbonate template until it reached the top edge,and degassed again to remove residual air bubbles. The PDMS was cured at65° C. for at least four hours and removed from the polycarbonate moldusing tweezers. A 1.5 mm biopsy punch was used to create inlet andoutlet channels leading to the inlet and outlet reservoirs in the PDMSslab. Barbed nylon connectors (inner diameter: 1/16″, McMaster-Carr,Elmhurst, Ill., USA) were inserted into the inlet and outlet channelsand sealed using PDMS to minimize leaking and provide more stabilityduring device reuse.

The top and bottom acrylic panels of the device were designed usingCorelDRAW ×7 (Corel Corporation, Ottawa, Canada) software. To match thedimensions of the PDMS slab, 1.5 mm thick acrylic sheets (ePlastics, SanDiego, Calif., USA) were cut into 20 mm×20 mm squares using an EpilogMini 18 (Epilog Laser, Golden, Colo., USA) at 23% speed, 50% power, and2500 Hz. Four holes with 3 mm diameter were cut in the panelsequidistant from the corners to accommodate bolts. For the top acrylicpanels, rectangular vents were cut above the inlet and outlet reservoirsto release gas bubbles collected by the polytetrafluoroethylene (PTFE)membrane, described below. The circular pieces from the holes andwindows were removed using tweezers and the acrylic pieces were cleanedwith sterile water to remove debris from the fabrication process. A0.005″ PTFE sheet (ePlastics, San Diego, Calif., USA) was laser-cut at12% speed, 10% power, and 2500 Hz into 20 mm×20 mm squares with voidslocated at the triangular compartments and bolt holes, matching theacrylic panels. PTFE membranes were brushed with uncured PDMS and bondedto the bottom surface of top acrylic panels for easier device assembly.

PDMS slabs were sandwiched between top and bottom acrylic panels and a2.5 mm stainless steel biopsy punch was manually fed through thecircular holes in the acrylic to create holes in the PDMS slab that wereregistered with the acrylic panels and PTFE membrane. At the time ofexperiments, a PDMS slab and PTFE membrane was clamped between a top andbottom acrylic panel using stainless steel, button head, Torx Plus bolts(thread size: 4-40, length: ⅜″, McMaster-Carr, Elmhurst, Ill., USA) andnarrow hex nuts (thread size: 4-40, height: 1/16″, McMaster-Carr,Elmhurst, Ill., USA).

Multiphysics Modeling

Multiphysics modeling was performed using the finite element softwarepackage, COMSOL Multiphysics 5.3 (COMSOL Group, Stockholm, Sweden). TheReynolds number was calculated to be 0.0566 using the fluid velocity,density and viscosity of the media, and the dimensions of the channels.The scale of our device falls within the laminar flow regime; therefore,flow can be analyzed using the “laminar flow physics interface (spf)”.The following assumptions were incorporated into the model: (i) thesurface condition was set to no-slip because of the solid boundaries;(ii) the fluid was assumed to have the same density and viscosity aswater; (iii) the inlet velocity was set to be a constant 0.25 mL/hr andthe outlet surface was treated as a free surface, allowing fluid to passthrough freely, (iv) the shear stress on the bounding surfaces wascalculated by multiplying shear rate (COMSOL variable, spf.sr) withdynamic viscosity (user defined, spf.mu). Studies were conducted instationary mode, as the conditions are steady-state. Velocity fields,streamlines, and surface shear stresses were plotted in various viewsand slices to show specific areas of interest, such as the center of theinlet and outlet channels and the surfaces of the hearts.

Zebrafish Heart Harvest and Culture for Fresh Hearts and Dish Hearts

Zebrafish lines Tg(fli1a:EGFP)^(y1) (26) and Tg(−0.8flt1:RFP)^(hu5333)(referred to as flt1^(enh):tdtomnato) (27) were raised and maintained atChildren's Hospital Los Angeles (CHLA) under standard conditions of careand with CHLA Institutional Animal Care and Use Committee (IACUC)oversight. IACUC vetted and provided prior approval for all experimentalprocedures used in this study. Zebrafish were euthanized with Tricainesolution (4.2 mL of Tricaine stock in 100 mL of fish water) andpositioned ventral side up in the groove of a moistened sponge under adissecting microscope. The chest wall was opened with a longitudinal cutbetween the gills. The heart was gently pulled out of the chest cavitywith a pair of tweezers holding the bulbus arteriosus and severed at theventral aorta and sinus venosus using microscissors. The hearts wererinsed in Ringer's solution (115 mM sodium chloride, 2.9 mM potassiumchloride, 1.8 mM calcium dichloride, 5 mM HEPES, pH 7.2 at 28° C.) threetimes before culturing. For Dish culture, 1 mL of pre-warmed media(DMEM, 10% FBS, 1:500 Primocin, 100 u⁻¹ Penicillin, 100 μg/mL Step, 2μg/mL Heparin) was placed in a 12-well cell culture plate, into whichone heart was placed per well and then statically incubated at 28.5° C.Media was replaced every second day using standard sterile culturetechniques.

Loading Hearts into Device

To culture hearts in the fluidic device, bolts were inserted through thefour holes in a bottom acrylic panel and PDMS slab to hold thesecomponents together. The PDMS slab was pre-wet with 70% ethanol tominimize trapping of air bubbles and then submerged in Ringer'ssolution. Any air bubbles attached to the PDMS were dislodged by manualpipetting. Extracted zebrafish hearts were placed in each compartmentwith part of the ventricle fitting into the apex of the triangle. Thetop acrylic piece with the PTFE membrane was then positioned on top ofthe PDMS slab and secured by twisting a nut onto each bolt. Tubing wasinserted onto the barbed connectors while the device was still submergedin Ringer's solution.

A programmable two-channel syringe pump (NE-4000, SyringePump.com,Suffolk, N.Y., USA) was used to perfuse media into the device through1/16″ ID, ⅛″ OD PVC clear tubing (Masterkleer 5233K51, McMaster-Carr,Elmhurst, Ill., USA). 60 mL syringes filled with pre-warmed maintenancemedia (DMEM, 10% FBS, 1:500 Primicin, 100 u⁻¹ Penicillin, 100 μg/mLStep, 2 μg/mL Heparin) were attached to tubing connected to the devicesand loaded onto the syringe pump. All experiments were conducted at aperfusion rate of 0.25 mL/hr.

Histology and Structural Characterization

Hearts from each condition were fixed and processed for paraffinsectioning and Acid Fuchsin Orange G (AFOG) histology, as previouslydescribed (8). Sections were imaged using an optical microscope (OlympusBX51, Olympus Corporation, Tokyo, Japan). Images viewed using ImageJwere scored on the following criteria: normal morphology (similar tofreshly isolated hearts) was considered Score 1, any morphologyrestricted to the outer (cortical) layer of cardiac tissue wasconsidered Score 2, and any abnormal morphology in the inner(trabecular) layer or extending into this layer was considered Score 3.

Calcium Imaging and Analysis

Hearts from each condition were transferred to Petri dishes andincubated in 10 μM Rhod-2 AM (R 1245MP, ThermoFisher Scientific,Waltham, Mass., USA) in maintenance media for 20 minutes at 28° C.Samples were then rinsed and imaged in Tyrode's solution (0.5 mM HEPES,0.1 mM magnesium chloride, 0.54 mM potassium chloride, 0.33 mM sodiumphosphate, 1.8 mM calcium chloride, 5.0 mM glucose, pH 7.4 at 37° C.)with 15 μM blebbistatin to arrest contraction and minimize motionartifacts during data collection. Each Petri dish was moved to the stageof an inverted fluorescent microscope (Nikon Eclipse Ti, NikonCorporation, Tokyo, Japan) enclosed in a 28° C. incubation chamber(OKOLAB USA Inc., San Bruno, Calif., USA). A platinum point electrodewas connected to a stimulator (MyoPacer MYP100, IonOptix, Westwood,Mass., USA) and positioned into the dish approximately 2-3 mm away fromthe hearts using a micro-manipulator (Patchman NP 2, Eppendorf,Hauppauge, N.Y., USA). The electrode was used to deliver 35 V biphasic,charge-balanced pulses at 0.5, 1.0, 1.5, and 2.0 Hz. Fluorescence of thecalcium transients was captured using a 10× air objective and ahigh-speed camera (Andor Zyla sCMOS, Oxford Instruments, Abingdon, UK)at 100 frames per second, 4×4 binning, and a gain of 4.

Calcium transients were extracted by plotting average fluorescentintensity of a 100×100 pixel region of interest using ImageJ. Thesevalues were then processed using custom MATLAB software (MATLAB,MathWorks, Natick, Mass., USA), similar to previous studies (40). Aftercalculating the period of the transients within each trace with aFourier transform, a peak detection algorithm was used to automaticallysplit each trace into individual transients. Each transient was analyzedto calculate rise time and decay times. The largest sustainedfluorescence increase was identified as the rising slope of thetransient. Backtracking in time until the slope changed from positive tonegative provided the start time and background fluorescence signal. Therise time was found by subtracting this start time from the time of themaximum intensity. To find the decay time, the background brightnessvalue was subtracted from the maximum brightness to find a normalizedmaximum brightness. Then, the time until this normalized brightnesssignal fell by 50% was calculated and defined as the decay time.

Live Imaging

Hearts were resected as previously described (8) and then isolated intoRinger's solution containing 100 μg/mL Primocin and 150 U/mL heparinfrom terminally anesthetized transgenic zebrafish after six dayspost-amputation. Isolated hearts were then transferred to wells of adevice and hooked up to the media perfusion system. The device wasperfused with media within the live imaging chamber of a fluorescentmicroscope (Leica DM IRE2, Leica Microsystems, Wetzlar, Germany) overfour to six days. Four hearts (in separate wells) were perfused in L-15media supplemented with 10% FCS, 100 μg/mL Primocin, 1.25 mM CaCl₂, and800 mg/L glucose at a rate of 0.3 mL/hr. μManager acquisition software(41) was used to capture a predefined z stack of images every hour forthe duration of the experiment. Acquired images were then concatenatedand cropped in ImageJ, and deconvolved using AutoQuant X3 (MediaCybernetics Inc, Rockville, Md.). Finally, focus and brightnessfluctuation were corrected using Gaussian Focus and BrightnessNormalizer plug-ins (supplemental downloads) in ImageJ.

Statistical Analysis

Statistical analysis was performed using GraphPad Prism 8 (GraphPadSoftware Inc., San Diego, Calif., USA). All data are plotted asindividual data points (biological replicates), with lines indicatingthe mean and error bars representing the standard deviation of the mean.A p value of less than 0.05 was considered statistically significant.All datasets were first tested for normality using the Shapiro-Wilktest. For histology scoring and capture rate measurements, data was notnormally distributed and thus the Kruskal-Wallis test followed by Dunn'smultiple comparisons test was used to compare the control group (Fresh)to each experimental group (one- and two-week Dish and Device hearts).For calcium transient rise and decay times, data was normallydistributed and thus two-way ANOVA followed by Tukey's multiplecomparisons test was used to compare the control group to eachexperimental group. For capture rate and calcium transient rise anddecay times, each frequency (0.5, 1.0, 1.5, and 2.0 Hz) wasstatistically compared independently.

While exemplary embodiments are described above, it is not intended thatthese embodiments describe all possible forms of the invention. Rather,the words used in the specification are words of description rather thanlimitation, and it is understood that various changes may be madewithout departing from the spirit and scope of the invention.Additionally, the features of various implementing embodiments may becombined to form further embodiments of the invention.

REFERENCES

-   1. M. A. Laflamme, C. E. Murry, Heart regeneration. Nature 473,    326-335 (2011).-   2. P. Staat, G. Rioufol, C. Piot, Y. Cottin, T. T. Cung, I.    L'Huillier, J. F. Aupetit, E. Bonnefoy, G. Finet, X. André-Fouët, M.    Ovize, Postconditioning the human heart. Circulation 112, 2143-2148    (2005).-   3. J. O. Oberpriller, J. C. Oberpriller, Response of the adult newt    ventricle to injury. J Exp Zool 187, 249-253 (1974).-   4. T. Borchardt, T. Braun, Cardiovascular regeneration in    non-mammalian model systems: what are the differences between newts    and man? Thromb Haemost 98, 311-318 (2007).-   5. J. Bloomekatz, M. Galvez-Santisteban, N. C. Chi, Myocardial    plasticity: cardiac development, regeneration and disease. Curr Opin    Genet Dev 40, 120-130 (2016).-   6. J. M. Gonzalez-Rosa, C. E. Burns, C. G. Burns, Zebrafish heart    regeneration: 15 years of discoveries. Regeneration (Oxf) 4, 105-123    (2017).-   7. N. Rubin, M. R. Harrison, M. Krainock, R. Kim, C. L. Lien, Recent    advancements in understanding endogenous heart regeneration-insights    from adult zebrafish and neonatal mice. Semin Cell Dev Biol 58,    34-40 (2016).-   8. K. D. Poss, L. G. Wilson, M. T. Keating, Heart regeneration in    zebrafish. Science 298, 2188-2190 (2002).-   9. K. Kikuchi, J. E. Holdway, A. A. Werdich, R. M. Anderson, Y.    Fang, G. F. Egnaczyk, T. Evans, C. A. Macrae, D. Y. Stainier, K. D.    Poss, Primary contribution to zebrafish heart regeneration by    gata4(+) cardiomyocytes. Nature 464, 601-605 (2010).-   10. C. Jopling, E. Sleep, M. Raya, M. Marti, A. Raya, J. C. Izpisua    Belmonte, Zebrafish heart regeneration occurs by cardiomyocyte    dedifferentiation and proliferation. Nature 464, 606-609 (2010).-   11. A. Lepilina, A. N. Coon, K. Kikuchi, J. E. Holdway, R. W.    Roberts, C. G. Burns, K. D. Poss, A dynamic epicardial injury    response supports progenitor cell activity during zebrafish heart    regeneration. Cell 127, 607-619 (2006).-   12. R. Marín-Juez, M. Marass, S. Gauvrit, A. Rossi, S. L. Lai, S. C.    Materna, B. L. Black, D. Y. Stainier, Fast revascularization of the    injured area is essential to support zebrafish heart regeneration.    Proc Natl Acad Sci USA 113, 11237-11242 (2016).-   13. M. Harrison, X. Feng, Q. Mo, A. Aguayo, J. Villafuerte, T.    Yoshida, C. Pearson, S. Schulte-Merker, C. Lien, Late developing    cardiac lymphatic vasculature supports adult zebrafish heart    function and regeneration. eLife. 14. W. C. Chen, Z. Wang, M. A.    Missinato, D. W. Park, D. W. Long, H. J. Liu, X. Zeng, N. A.    Yates, K. Kim, Y. Wang, Decellularized zebrafish cardiac    extracellular matrix induces mammalian heart regeneration. Sci Adv    2, e1600844 (2016).-   15. J. Kim, N. Rubin, Y. Huang, T. L. Tuan, C. L. Lien, In vitro    culture of epicardial cells from adult zebrafish heart on a fibrin    matrix. Nat Protoc 7, 247-255 (2012).-   16. V. Sander, G. Suñe, C. Jopling, C. Morera, J. C. Izpisua    Belmonte, Isolation and in vitro culture of primary cardiomyocytes    from adult zebrafish hearts. Nat Protoc 8, 800-809 (2013).-   17. C. L. Lien, M. Schebesta, S. Makino, G. J. Weber, M. T. Keating,    Gene expression analysis of zebrafish heart regeneration. PLoS    biology 4, e260 (2006).-   18. S. Pieperhoff, K. S. Wilson, J. Baily, K. de Mora, S.    Maqsood, S. Vass, J. Taylor, J. Del-Pozo, C. A. MacRae, J. J.    Mullins, M. A. Denvir, Heart on a plate: histological and functional    assessment of isolated adult zebrafish hearts maintained in culture.    PloS one 9, e96771 (2014).-   19. J. Cao, K. D. Poss, Explant culture of adult zebrafish hearts    for epicardial regeneration studies. Nat Protoc 11, 872-881 (2016).-   20. J. Akagi, K. Khoshmanesh, B. Evans, C. J. Hall, K. E.    Crosier, J. M. Cooper, P. S. Crosier, D. Wlodkowic, Miniaturized    embryo array for automated trapping, immobilization and    microperfusion of zebrafish embryos. PLoS One 7, e36630 (2012).-   21. L. L. Bischel, B. R. Mader, J. M. Green, A. Huttenlocher, D. J.    Beebe, Zebrafish Entrapment By Restriction Array (ZEBRA) device: a    low-cost, agarose-free zebrafish mounting technique for automated    imaging. Lab Chip 13, 1732-1736 (2013).-   22. M. Erickstad, L. A. Hale, S. H. Chalasani, A. Groisman, A    microfluidic system for studying the behavior of zebrafish larvae    under acute hypoxia. Lab Chip 15, 857-866 (2015).-   23. C. Liu, J. A. Thompson, H. H. Bau, A membrane-based,    high-efficiency, microfluidic debubbler. Lab Chip 11, 1688-1693    (2011).-   24. V. Vedula, J. Lee, H. Xu, C. J. Kuo, T. K. Hsiai, A. L. Marsden,    A method to quantify mechanobiologic forces during zebrafish cardiac    development using 4-D light sheet imaging and computational    modeling. PLoS Comput Biol 13, e1005828 (2017).-   25. F. Boselli, J. Vermot, Live imaging and modeling for shear    stress quantification in the embryonic zebrafish heart. Methods 94,    129-134 (2016).-   26. N. D. Lawson, B. M. Weinstein, In vivo imaging of embryonic    vascular development using transgenic zebrafish. Developmental    biology 248, 307-318 (2002).-   27. J. Bussmann, F. L. Bos, A. Urasaki, K. Kawakami, H. J.    Duckers, S. Schulte-Merker, Arteries provide essential guidance cues    for lymphatic endothelial cells in the zebrafish trunk. Development    137, 2653-2657 (2010).-   28. L. Zhao, A. L. Borikova, R. Ben-Yair, B. Guner-Ataman, C. A.    MacRae, R. T. Lee, C. G. Burns, C. E. Burns, Notch signaling    regulates cardiomyocyte proliferation during zebrafish heart    regeneration. Proc Natl Acad Sci USA 111, 1403-1408 (2014).-   29. E. Lin, S. Shafaattalab, J. Gill, B. Al-Zeer, C. Craig, M.    Lamothe, K. Rayani, M. Gunawan, A. Y. Li, L. Hove-Madsen, G. F.    Tibbits, Physiological phenotyping of the adult zebrafish heart. Mar    Genomics, 100701 (2019).-   30. E. R. Porrello, A. I. Mahmoud, E. Simpson, J. A. Hill, J. A.    Richardson, E. N. Olson, H. A. Sadek, Transient regenerative    potential of the neonatal mouse heart. Science 331, 1078-1080    (2011).-   31. M. A. Lancaster, J. A. Knoblich, Organogenesis in a dish:    modeling development and disease using organoid technologies.    Science 345, 1247125 (2014).-   32. D. Dutta, I. Heo, H. Clevers, Disease Modeling in Stem    Cell-Derived 3D Organoid Systems. Trends Mol Med 23, 393-410 (2017).-   33. C. A. MacRae, R. T. Peterson, Zebrafish as tools for drug    discovery. Nature reviews. Drug discovery 14, 721-731 (2015).-   34. C. A. Macrae, Cardiac Arrhythmia: In vivo screening in the    zebrafish to overcome complexity in drug discovery. Expert Opin Drug    Discov 5, 619-632 (2010).-   35. M. Vornanen, M. Hassinen, Zebrafish heart as a model for human    cardiac electrophysiology. Channels (Austin) 10, 101-110 (2016).-   36. D. J. Milan, I. L. Jones, P. T. Ellinor, C. A. MacRae, In vivo    recording of adult zebrafish electrocardiogram and assessment of    drug-induced QT prolongation. Am J Physiol Heart Circ Physiol 291,    H269-273 (2006).-   37. R. Arnaout, T. Ferrer, J. Huisken, K. Spitzer, D. Y.    Stainier, M. Tristani-Firouzi, N. C. Chi, Zebrafish model for human    long QT syndrome. Proceedings of the National Academy of Sciences of    the United States of America 104, 11316-11321 (2007).-   38. A. Pott, W. Rottbauer, S. Just, Functional genomics in zebrafish    as a tool to identify novel antiarrhythmic targets. Current    medicinal chemistry 21, 1320-1329 (2014).-   39. D. P. Yen, Y. Ando, K. Shen, A cost-effective micromilling    platform for rapid prototyping of microdevices. Technology (Singap    World Sci) 4, 234-239 (2016).-   40. A. P. Petersen, D. M. Lyra-Leite, N. R. Ariyasinghe, N.    Cho, C. M. Goodwin, J. Y. Kim, M. L. McCain, Microenvironmental    Modulation of Calcium Wave Propagation Velocity in Engineered    Cardiac Tissues. Cellular and molecular bioengineering, (2018).-   41. A. Edelstein, N. Amodaj, K. Hoover, R. Vale, N. Stuurman,    Computer control of microscopes using μManager. Curr Protoc Mol Biol    Chapter 14, Unit14.20 (2010).

What is claimed is:
 1. A microfluidic assembly comprising: a polymericblock having a top surface and a bottom surface, the polymeric blockdefining: an input reservoir; at least one triangular well; an outputreservoir; a first flow channel system in fluid communication with theinput reservoir and the at least one triangular well; a second flowchannel system in fluid communication with the output reservoir and theat least one triangular well, wherein the first flow channel system andthe second flow channel system each independently include tracks havinga closed bottom and open top; an inlet conduit in fluid communicationwith the input reservoir; an outlet conduit in fluid communication withthe output reservoir; a first transparent plate disposed over the topsurface of the polymeric block; and a second transparent plate disposedover the bottom surface of the polymeric block.
 2. The microfluidicassembly of claim 1 wherein the at least one triangular well that trapsand maintains at orientation of an explanted organ or organoid.
 3. Themicrofluidic assembly of claim 1 wherein the at least one triangularwell that traps and maintains at orientation of an explanted zebrafishheart.
 4. The microfluidic assembly of claim 1 wherein the at least onetriangular well that traps and maintains at orientation of mouseembryos/organs and human fetal organs.
 5. The microfluidic assembly ofclaim 1 further comprising a gas permeable membrane interposed betweenthe first transparent plate and the polymeric block.
 6. The microfluidicassembly of claim 5 wherein the gas permeable membrane has openings thatalign to triangular wells to allow imaging of organs placed therein. 7.The microfluidic assembly of claim 5 wherein the first transparent platedefines a first vent slot and a second vent slot allowing gases to ventfrom the gas permeable membrane, the first vent slot overlaying theinput reservoir and a portion of the first flow channel system, and thesecond vent slot overlaying the output reservoir and a portion of thesecond flow channel system.
 8. The microfluidic assembly of claim 5wherein the gas permeable membrane is a PTFE membrane.
 9. Themicrofluidic assembly of claim 1 wherein the polymer block comprisespolydimethylsiloxane.
 10. The microfluidic assembly of claim 1 whereinthe at least one triangular well includes a plurality of triangularwells.
 11. The microfluidic assembly of claim 1 wherein the at least onetriangular well includes 2 to 10 triangular wells.
 12. The microfluidicassembly of claim 1 wherein the polymeric block has a thickness fromabout 1 to 10 mm.